Electron microscopy work at Cheng laboratory is partly support by grants from NIH (1R01GM082893, 1R01GM098672, 1S10RR026814-01 and P50GM082250 (to A. Frankel)), and a grant from UCSF Program of Breakthrough Biomedical Research, Opportunity Award and New Technology Award.

1. Preparation of EM grids for carbon coating
<ol>
 <li>Fill a large glass beaker with distilled water.</li>
 <li>Use a glass pipette to drop a single drop of collodion (~20&mu;l, 2% in amyl acetate, Electron Microscopy Sciences, PA, USA) on the surface of the water. Collodion will spread to form a thin plastic layer at the water/air surface; Note: A first drop of collodion may be used to clear the water surface from any dust particles. After removing of the film from the first drop, the water surface is clear of dust.</li>
 <li>Place EM grids, 400 or 200 mesh Gilder Cu grids purchased from Ted Pella Inc. (USA), one by one onto the plastic layer floating on the water surface; We usually place 50 ~ 100 grids in a close packing pattern, with the shiny side of the grid facing down into the plastic layer (Fig. 1A). Note: 400 mesh grids are good for routine negative stain EM grids; whereas 200 mesh grids are good for collecting tilt pair images.</li>
 <li>Cut a piece of paper with dimensions slightly larger than the area where grids are placed on the plastic layer. Place the paper carefully on top of the grid (Fig. 1B), and wait until the paper becomes completely wet (Fig. 1C). Note: Different types of paper have different water absorption rates. It is important that the paper does not absorb water too fast. For example, regular filter paper is not good for this purpose because it absorbs water too fast and makes a lot of wrinkles. The carbon film made from wrinkled paper is not flat, and thus not good for collecting tilt pair images for random conical tilt 3D reconstruction. One has to test different papers to find a suitable one. In our laboratory, we have been using papers from notepads bought from local grocery store. Sometimes the newspaper paper was also found to be suitable. Another option is to use the relative thick rice paper used for Chinese/Japanese calligraphy.</li>
 <li>Rapidly make incisions on the plastic film around and close to the soaked paper. Immediately afterwards, use a pair of forceps to pick up the paper with the grids sandwiched between the plastic film and paper. Place the paper in a petri dish with the grids facing up for air drying (Fig. 1D); Note: It is important to let the paper air-dry. Drying too fast generates many wrinkles in the paper.</li>
</ol>
2. Carbon coating grids
<ol>
 <li>The carbon-coating machine used in our laboratory is a Cressington 208C High Vacuum Turbo Carbon Coater (Ted Pella). Our system is equipped with a Cressington film thickness monitor, which measures the thickness of coated carbon film based on the principle that the oscillating frequency of a quartz crystal is changed by the mass of a deposited film on its surface. However, the protocol described here does not require a thickness monitor, since it is often not available in every laboratory.</li>
 <li>Sharpen a carbon rod using a sharpener (Ted Pella) and load carbon rods in carbon coating machine (Fig. 2A). Two rods, one with a sharp tip and the other with a flat end are placed against each other.</li>
 <li>Applying vacuum grease on half of the frosted area of a glass slide and place it next to the grids (Fig. 2B). Only half of the frosted area of the glass slide should be covered with vacuum grease. Note: The area with vacuum grease will not change color after carbon coating. The contrast between the adjacent areas with and without vacuum grease provides an estimation of thickness of coated carbon.</li>
 <li>Place the paper with grids on the stage of carbon coating machine. Place the glass slide next to the grids (Fig. 2C). Note: One can also first transfer grids coated with the plastic film to a glass slide before coating, in case many grids fall off from the paper.</li>
 <li>Pump the vacuum chamber, and wait until the vacuum reaches a level of 10-5 Torr. Note: coating with lower vacuum often generates many sparks, resulting to many large carbon particles on the grid.</li>
 <li>Increase the current very slowly. The sharp tip of the carbon rod will first become red and then white as the current increases. Do not watch the high brightness tip directly with the naked eye. Stop increasing current when the coating starts. The glass slide will change color from white to light grey and then dark grey. The darkness of the slides correlates the thickness of the carbon film being coated. Turn the current off to stop coating when desired thickness has been achieved (Fig. 2D). Note: 1) After repeated carbon coating processes, coated carbon from the inside of the glass bell jar could darken its color. This makes the coated carbon film appear thicker than it actually is. 2) One sharpened carbon rod can be used for several coating. The sharpness of the tip is gradually reduced after each coating. The maximum current is influenced by the sharpness of the carbon rod. A freshly sharpened rod with smaller diameter requires a relative lower current to evaporate carbon. Later coating may require slightly higher current.</li>
 <li>Vent the vacuum chamber and remove coated grids for later use. 
 Note 1. The more accurate method of controlling the carbon thickness is by using a film thickness monitor. Or one can qualitatively calibrate color against the accurate thickness measured by the monitor. A useful carbon thickness for this technique is between 3 and 7 nm.</li>
</ol>
This is a simple and easy method to prepare a large quantity of carbon coated grids, usually ~100 grids per preparation. The grids contain a thin plastic layer, which usually does not affect the quality of the negatively stained image. However, grids prepared in this method are not suitable for cryoEM. This layer of plastic can be removed by soaking the grids into chloroform for several hours.
3. Preparing uranyl formate solution (0.75%) for negative stain EM. The detailed protocol was described previously 5
<ol>
 <li>Weigh out 37.5mg of uranyl formate into a small beaker;</li>
 <li>Add 5 ml of boiling water and stir for 5 minutes in the dark;</li>
 <li>Add 10&mu;l 5M NaOH, continue stir for 5 minutes;</li>
 <li>Filter the solution using a syringe filter (Fisherbrand, 13mm syringe filter, 0.2&mu;m, Nylon non-sterile, catalog number 09-720-5), and store the filtered solution in a 8ml Falcon tube covered with aluminum foil; 
 Note. Uranyl Formate powder can be purchased from Electron Microscopy Sciences (Hatfield, PA), catalog number 22451. Uranyl formate solution is light sensitive and should be kept away from direct light, for example by covering the Falcon tube with aluminum foil. A freshly prepared solution has a light yellow color (Fig. 3A). Stain solution can be kept on the bench for one to two weeks before it starts precipitating. Uranyl formate is acidic with a pH ~4. Adding NaOH increases the pH, but adding it too quickly or in excess can cause the stain to precipitate. In addition to generating contrast, uranyl formate also fixes the protein sample. Because the fixation rate is so rapid, the low pH usually does not affect the overall protein conformations or protein complex formation 7.</li>
</ol>
4. Protocol for preparing negative stain EM grid. This protocol was described in detail previously 5
<ol>
 <li>Glow-discharge carbon coated grid, using PELCO easiGlow GlowDischarge system (Ted Pella Inc., USA). The current used is 15 mA and grids are glow discharged for 30 sec. Note. Glow-discharged grids should be used shortly after glow discharging. We often use the grids in less than 2 hours after the glow discharge.</li>
 <li>Place two 20&mu;l drops of distilled water and two 25&mu;l drops of uranyl formate solution on a piece of parafilm (Fig. 3B).</li>
 <li>Hold a glow-discharged grid using reverse-force anti-capillary forceps with the carbon-coated side facing up. Note: It is necessary to use anti-capillary forceps. Otherwise, liquid sample will be sucked away from grids by the capillary force between the forceps.</li>
 <li>Apply 2&mu;l protein sample to the glow-discharged grid, and wait for 30 sec. Note: The waiting time and the length of glow discharge influence the concentration of proteins adsorbed in the grid. Very diluted samples require longer adsorption times. However, prolonged adsorption may change the buffer concentration since water vapors. To achieve a proper particle distribution in grid requires somewhat trial and error approach.</li>
 <li>Blot off the sample using filter paper.</li>
 <li>Touch the grid surface to a water drop and then blot off the water with filter paper. Repeat with the 2nd drop of water.</li>
 <li>Touch the grid surface to the first drop of stain briefly. Blot off the stain solution with filter paper.</li>
 <li>Touch the grid surface to the second drop of stain for 20 seconds. Blot off stain solution with filter paper.</li>
 <li>Dry the grid using a vacuum. Note: one can also leave grids on the bench or in a chemical hood to air dry. The benefit of vacuum dry is that one can observe the grids in the EM immediately. The possible artifact generated by quick drying is uneven stain.</li>
</ol>
5. Transmission electron microscope
<ol>
 <li>The microscope used in this demonstration is a Tecnai T12 (FEI Company) equipped with a Gatan 4K x 4K CCD camera.</li>
 <li>Briefly check the alignment of the microscope before inserting sample; Note: This is only needed once per EM session. In a model microscope, near perfect alignment can be saved earlier. Loading such an alignment file should restore the microscope alignment to a near perfect condition.</li>
 <li>Load the EM grid in the single tilt standard specimen holder.</li>
 <li>Insert holder into CompuStage of the T12. Wait for the pumping cycle to complete.</li>
 <li>Insert the holder into column.</li>
 <li>Focus specimen and set a desired defocus, usually -1.5um. Note: Focusing can be achieved either by manually focusing using the minimum contrast method or using CCD camera to capture live images and compute live Fourier power spectrum.</li>
 <li>Record an image using CCD camera;</li>
 <li>In images of negatively stained proteins samples, proteins are often seen with white contrast, i.e. white density against a darker background. Note that while the protein concentrations in different areas of the same grid are usually quite similar, one does observe different levels of stain in different areas of the same grid. Sometimes areas could have uneven staining or even positive staining (protein appears dark on a light background), whereas some other areas have perfect stain. It is often necessary to search the grid to locate a good area for imaging.</li>
</ol>
6. Representative Results
Examples of typical good images of negatively stained samples, horse spleen ferritin (Figure 4A), Fab (Fig. 4B), archaeal 20S proteasome (Fig. 4C) and nucleosome (Fig. 4D), are shown.
<img alt="Figure 1" src="/files/ftp_upload/3227/3227fig1.jpg" /> 
Figure 1. Preparation of carbon coated grids for negative stain EM. A) Placing EM grids one-by-one in a closed packing pattern on the surface of collodion film. B) Placing a piece of paper slightly larger than the area of the grids. C) Wait until the paper gets completely wet. D) Pick up the paper together with the grids for air-drying.
<img alt="Figure 2" src="/files/ftp_upload/3227/3227fig2.jpg" /> 
Figure 2. Coating EM grids with carbon. A) Setup the carbon evaporator: place the two carbon rods in the coating apparatus, with one rod (on the left) sharpened to a point tip. The other rod (on the right) has a flat surface and come in contact with the point tip of the one in the opposite side. B) Place a small amount of vacuum grease in one side of a fostered glass slide. The area within the black frame has vacuum grease. C) Setup the grids for coating. The glass slides are used to hold the paper in position. The arrowhead points the area with vacuum grease. D) After coating, the color of the slide with vacuum grease covered does not change the color. The contrast indicates the thickness of coated carbon.
<img alt="Figure 3" src="/files/ftp_upload/3227/3227fig3.jpg" /> 
Figure 3. Illustrating the negative staining procedure. A) Freshly prepared (left) uranyl formate solution is transparent without sign of precipitates. After 1~2 weeks, precipitates start accumulating in the bottom in the bottom of the tube. B) Place two drops (~25&mu;l) of distilled water and two drops of uranyl formate solution in a piece of parafilm. Use a pair of anti-capillary forceps to hold a glow discharged carbon-coated EM grid with carbon side up. A drop of sample (~2&mu;l) is placed in the grid. C) After waiting for 30 second, blot the solution with a filter paper. D) Flip the grid to touch the sample side of the grid with the first drop of water. Repeat the step C and D to complete the process of staining.
<img alt="Figure 4" src="/files/ftp_upload/3227/3227fig4.jpg" /> 
Figure 4. Examples of EM images of negatively stained protein samples. A) Horse spleen ferritin, B) fragment of antigen binding (Fab), C) archaeal 20S proteasome and D) nucleosome. All images were recorded with the same magnification. The scale bar is 20nm.

<ol>
	<li>Cheng, Y., Walz, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+advent+of+near-atomic+resolution+in+single-particle+electron+microscopy.">The advent of near-atomic resolution in single-particle electron microscopy.</a> Annu. Rev. Biochem. 78, 723-742 (2009).</li><li>Grigorieff, N., Harrison, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Near-atomic+resolution+reconstructions+of+icosahedral+viruses+from+electron+cryo-microscopy.">Near-atomic resolution reconstructions of icosahedral viruses from electron cryo-microscopy.</a> Curr. Opin. Struct. Biol. 21 (2), 265-273 (2011).</li><li>Zhou, Z. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+resolution+cryo+electron+microscopy+of+macromolecular+complexes.">Atomic resolution cryo electron microscopy of macromolecular complexes.</a> Adv. Protein Chem. Struct. Biol. 82, 1-35 (2011).</li><li>Adrian, M., Dubochet, J., Lepault, J., McDowall, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryo-electron+microscopy+of+viruses.">Cryo-electron microscopy of viruses.</a> Nature. 308 (5954), 32-36 (1984).</li><li>Ohi, M., Li, Y., Cheng, Y., Walz, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Negative+Staining+and+Image+Classification+-+Powerful+Tools+in+Modern+Electron+Microscopy.">Negative Staining and Image Classification - Powerful Tools in Modern Electron Microscopy.</a> Biol. Proced. Online. 6, 23-34 (2004).</li><li>Rabl, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanism+of+gate+opening+in+the+20S+proteasome+by+the+proteasomal+ATPases.">Mechanism of gate opening in the 20S proteasome by the proteasomal ATPases.</a> Mol. Cell. 30 (3), 360-368 (2008).</li><li>Zhao, F. Q., Craig, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capturing+time-resolved+changes+in+molecular+structure+by+negative+staining.">Capturing time-resolved changes in molecular structure by negative staining.</a> J. Struct. Biol. 141 (1), 43-52 (2003).</li><li>Radermacher, M., Wagenknecht, T., Verschoor, A., Frank, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+reconstruction+from+a+single-exposure,+random+conical+tilt+series+applied+to+the+50S+ribosomal+subunit+of+Escherichia+coli.">Three-dimensional reconstruction from a single-exposure, random conical tilt series applied to the 50S ribosomal subunit of Escherichia coli.</a> J. Microsc. 146 (Pt. 2), 113-136 (1987).</li><li>Zhao, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-dimensional+crystallization+conditions+of+human+leukotriene+C4+synthase+requiring+adjustment+of+a+particularly+large+combination+of+specific+parameters.">Two-dimensional crystallization conditions of human leukotriene C4 synthase requiring adjustment of a particularly large combination of specific parameters.</a> J. Struct. Biol. 169 (3), 450-454 (2010).</li><li>Nishida, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+leukocyte+beta2+integrins+by+conversion+from+bent+to+extended+conformations.">Activation of leukocyte beta2 integrins by conversion from bent to extended conformations.</a> Immunity. 25 (4), 583-594 (2006).</li></ol>

Production and Free Access of this video-article is sponsored by FEI.

Negative stain EM is a powerful tool that can be used to study 3D structures of purified protein samples, such as using random conical tilt 8 method, which requires collecting a pair of images with one from specimen tilted to 60&deg; and one from the same area but untilted, to calculate 3D reconstructions of macromolecular complexes. In addition, negative staining EM can have many other applications, such as screening for two-dimensional (2D) crystals of membrane proteins 9, visualizing protein complexes in different conformations 10, or simply for visualizing homogeneity and/or heterogeneity of purified proteins sample thus providing feedbacks to improve protein purification procedures, etc. It can be a powerful assay for biochemists. In this article, we presented the protocols ranging from preparing the carbon-coated grids to collecting images of negatively stained protein samples in an electron microscope. As illustrated here, the procedure is rather simple and easy to follow. If attempting to calculate 3D reconstructions of the sample being observed by random conical tilt method, it is necessary to acquire tilt pair images. However, if performing single particle analysis is the purpose, multiple untilted images will suffice. In both cases, further image processing is required and this is beyond the scope of this video article.
As with any method, there are variations to the protocols. Uranyl formate works very well as a negative stain reagent in most cases. Uranyl acetate is another commonly used negative stain reagent. It works very similar as uranyl formate, but with a larger grain size and, in many cases, lower contrast than uranyl formate. The advantage of using uranyl acetate is that the solution lasts longer than uranyl formate. One does not need to prepare a fresh solution every one or two weeks. Both uranyl formate and acetate are acidic. An obvious concern is that the low pH could induce changes in the protein sample being studied. Both stain solutions also function as protein fixatives. An earlier study has shown that both fix protein samples quickly 7. This rapid fixation makes the low pH less important. During the process of staining, a protein sample becomes fixed before the low pH can induce any changes. Nevertheless, other negative stain reagents, such as ammonium molybdate and phosphotungstic acid, may yield better results for staining samples when low pH is a concern 7. Both ammonium molybdate and phosphotungstic acid have an approximately neutral pH, but the contrast of protein samples generated by these stains is usually inferior to uranyl formate.

<table border="1">
<tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalog 			Number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Collodion</td>
 <td>Electron 			Microscopy Sciences, PA, USA</td>
 <td>12620-30</td>
 <td>30ml, 2% in 			Amyl Acetate</td>
 </tr>
 <tr>
 <td>Gilder Cu 			grids</td>
 <td>Ted Pella 			Inc., USA</td>
 <td>G400/G200</td>
 <td>400/200 mesh</td>
 </tr>
 <tr>
 <td>Cressington 			208C High Vaccume Turbo Carbon Coater</td>
 <td>Ted Pella 			Inc., USA</td>
 <td>9260</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Double pointed 			Carbon rod</td>
 <td>Ted Pella 			Inc., USA</td>
 <td>58</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Uranyl formate</td>
 <td>Electron 			Microscopy Sciences, PA, USA</td>
 <td>16984-59-1</td>
 <td>99.9% purity</td>
 </tr>
 <tr>
 <td>PELCO easiGlow 			GlowDischarge system</td>
 <td>Ted Pella 			Inc., USA</td>
 <td>91000</td>
 <td>Purchase at 			local pharmacy</td>
 </tr>
 <tr>
 <td>Whatman #1 			Filter paper</td>
 <td>Fisher 			Scientific</td>
 <td>09-805E</td>
 <td>&nbsp;
 </td>
 </tr>
 </tbody>
</table>

visualizing proteins and macromolecular complexes by negative stain em: from grid preparation to image acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Watch this Scientific Journal Video about Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition at JoVE.com

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

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58.6K ビュー.  University of California San Francisco. 負染色電子顕微鏡（EM）で視覚化するタンパク質サンプルは、ポピュラーな構造解析の方法となっている。それは、タンパク質製剤の品質の定性試験のためにもそのような研究されている分子の三次元再構成を計算すると、定量的な構造解析に有用である、と。この記事では、EMグリッドを準備する試料を染色し、電子顕微鏡で試料を可視化するための詳?...

ビデオ: Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Cell Co-culture Patterning Using Aqueous Two-phase Systems

Cell patterning technologies that are fast, easy to use and affordable will be required for the future development of high throughput cell assays, platforms for studying cell-cell interactions and tissue engineered systems. This detailed protocol describes a method for generating co-cultures of cells using biocompatible solutions of dextran (DEX) and polyethylene glycol (PEG) that phase-separate when combined above threshold concentrations. Cells can be patterned in a variety of configurations using this method. Cell exclusion patterning can be performed by printing droplets of DEX on a substrate and covering them with a solution of PEG containing cells. The interfacial tension formed between the two polymer solutions causes cells to fall around the outside of the DEX droplet and form a circular clearing that can be used for migration assays. Cell islands can be patterned by dispensing a cell-rich DEX phase into a PEG solution or by covering the DEX droplet with a solution of PEG. Co-cultures can be formed directly by combining cell exclusion with DEX island patterning. These methods are compatible with a variety of liquid handling approaches, including manual micropipetting, and can be used with virtually any adherent cell type.

Aqueous two-phase systems were used to simultaneously pattern multiple populations of cells. This fast and easy method for cell patterning takes advantage of the phase separation of aqueous solutions of dextran and polyethylene glycol and the interfacial tension that exists between the two polymer solutions.

Cell patterning  technologies that are fast, easy to use and affordable will be required for the  future development of high throughput cell assays, ...

Preparation of Hydroxy-PAAm Hydrogels for Decoupling the Effects of Mechanotransduction Cues

It is now well established that many cellular functions are regulated by interactions of cells with physicochemical and mechanical cues of their extracellular matrix (ECM) environment. Eukaryotic cells constantly sense their local microenvironment through surface mechanosensors to transduce physical changes of ECM into biochemical signals, and integrate these signals to achieve specific changes in gene expression. Interestingly, physicochemical and mechanical parameters of the ECM can couple with each other to regulate cell fate. Therefore, a key to understanding mechanotransduction is to decouple the relative contribution of ECM cues on cellular functions.
Here we present a detailed experimental protocol to rapidly and easily generate biologically relevant hydrogels for the independent tuning of mechanotransduction cues in vitro. We chemically modified polyacrylamide hydrogels (PAAm) to surmount their intrinsically non-adhesive properties by incorporating hydroxyl-functionalized acrylamide monomers during the polymerization. We obtained a novel PAAm hydrogel, called hydroxy-PAAm, which permits immobilization of any desired nature of ECM proteins. The combination of hydroxy-PAAm hydrogels with microcontact printing allows to independently control the morphology of single-cells, the matrix stiffness, the nature and the density of ECM proteins. We provide a simple and rapid method that can be set up in every biology lab to study in vitro cell mechanotransduction processes. We validate this novel two-dimensional platform by conducting experiments on endothelial cells that demonstrate a mechanical coupling between ECM stiffness and the nucleus.

We present a new polyacrylamide hydrogel, called hydroxy-PAAm, that allows a direct binding of ECM proteins with minimal cost or expertise. The combination of hydroxy-PAAm hydrogels with microcontact printing facilitates independent control of many cues of the natural cell microenvironment for studying cellular mechanostransduction.

It is now well established that many cellular functions are regulated by interactions of cells with physicochemical and mechanical cues of their ...

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern tissue engineering. A technical bottleneck is the deposition of collagen in vitro, as it is notoriously slow, resulting in sub-optimal formation of connective tissue and subsequent tissue cohesion, particularly in skin models. Here, we describe a method which involves the addition of differentially-sized sucrose co-polymers to skin cultures to generate macromolecular crowding (MMC), which results in a dramatic enhancement of collagen deposition. Particularly, dermal fibroblasts deposited a significant amount of collagen I/IV/VII and fibronectin under MMC in comparison to controls.
The protocol also describes a method to decellularize crowded cell layers, exposing significant amounts of extracellular matrix (ECM) which were retained on the culture surface as evidenced by immunocytochemistry. Total matrix mass and distribution pattern was studied using interference reflection microscopy. Interestingly, fibroblasts, keratinocytes and co-cultures produced cell-derived matrices (CDM) of varying composition and morphology. CDM could be used as &#34;bio-scaffolds&#34; for secondary cell seeding, where the current use of coatings or scaffolds, typically from xenogenic animal sources, can be avoided, thus moving towards more clinically relevant applications.
In addition, this protocol describes the application of MMC during the submerged phase of a 3D-organotypic skin co-culture model which was sufficient to enhance ECM deposition in the dermo-epidermal junction (DEJ), in particular, collagen VII, the major component of anchoring fibrils. Electron microscopy confirmed the presence of anchoring fibrils in cultures developed with MMC, as compared to controls. This is significant as anchoring fibrils tether the dermis to the epidermis, hence, having a pre-formed mature DEJ may benefit skin graft recipients in terms of graft stability and overall wound healing. Furthermore, culture time was condensed from 5 weeks to 3 weeks to obtain a mature construct, when using MMC, reducing costs.

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

We present a protocol to obtain cell-derived matrices rich in extracellular matrix proteins, using macromolecular crowders (MMC). In addition, we present a protocol which incorporates MMC in 3D organotypic skin co-culture generation, which reduces culture time while maintaining maturity of construct.

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern ...

3D Magnetic Stem Cell Aggregation and Bioreactor Maturation for Cartilage Regeneration

Cartilage engineering remains a challenge due to the difficulties in creating an in vitro functional implant similar to the native tissue. An approach recently explored for the development of autologous replacements involves the differentiation of stem cells into chondrocytes. To initiate this chondrogenesis, a degree of compaction of the stem cells is required; hence, we demonstrated the feasibility of magnetically condensing cells, both within thick scaffolds and scaffold-free, using miniaturized magnetic field sources as cell attractors. This magnetic approach was also used to guide aggregate fusion and to build scaffold-free, organized, three-dimensional (3D) tissues several millimeters in size. In addition to having an enhanced size, the tissue formed by magnetic-driven fusion presented a significant increase in the expression of collagen II, and a similar trend was observed for aggrecan expression. As the native cartilage was subjected to forces that influenced its 3D structure, dynamic maturation was also performed. A bioreactor that provides mechanical stimuli was used to culture the magnetically seeded scaffolds over a 21-day period. Bioreactor maturation largely improved chondrogenesis into the cellularized scaffolds; the extracellular matrix obtained under these conditions was rich in collagen II and aggrecan. This work outlines the innovative potential of magnetic condensation of labeled stem cells and dynamic maturation in a bioreactor for improved chondrogenic differentiation, both scaffold-free and within polysaccharide scaffolds.

Chondrogenesis from stem cells requires fine tuning the culture conditions. Here, we present a magnetic approach for condensing cells, an essential step to initiate chondrogenesis. In addition, we show that dynamic maturation in a bioreactor applies mechanical stimulation to the cellular constructs and enhances cartilaginous extracellular matrix production.

Cartilage engineering remains a challenge due to the difficulties in creating an in vitro functional implant similar to the native tissue. An approach ...

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause ...

Preparation and Characterization of Nanoliposomes for the Entrapment of Bioactive Hydrophilic Globular Proteins

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their biocompatibility, biodegradability, non-immunogenicity, non-toxicity, and ability to entrap both hydrophilic and hydrophobic compounds. The classical hydration of thin lipid films in an organic solvent is applied herein as a technique to encapsulate tarin, a plant lectin, in nanoliposomes. Nanoliposome size, stability, entrapment efficiency, and morphological characterization are described in detail. The nanoliposomes are prepared using 1,2-dioleoyl-sn-glycerol-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (ammonium salt; DSPE-MPEG 2000), and cholesterylhemisuccinate (CHEMS) as the main constituents. Lipids are first dissolved in chloroform to obtain a thin lipid film that is subsequently rehydrated in ammonium sulfate solution containing the protein to be entrapped and incubated overnight. Then, sonication and extrusion techniques are applied to generate nanosized unilamellar vesicles. The size and polydispersity index of the nanovesicles are determined by dynamic light scattering, while nanovesicle morphology is assessed by scanning electron microscopy. Entrapment efficiency is determined by the ratio of the amount of unencapsulated protein to original amount of initially loaded protein. Homogeneous liposomes are obtained with an average size of 155 nm and polydispersity index value of 0.168. A high entrapment efficiency of 83% is achieved.

This study describes classical hydration using the thin lipid film method for nanoliposome preparation followed by nanoparticle characterization. A 47 kDa-hydrophilic and globular protein, tarin, is successfully encapsulated as a strategy to improve stability, avoid fast clearance, and promote controlled release. The method can be adapted to hydrophobic molecules encapsulation.

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their ...

In Vitro Model of Human Cutaneous Hypertrophic Scarring using Macromolecular Crowding

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol applies a macromolecular crowding (MMC) technique to mimic this physiological crowding through the addition of neutral polymers (crowders) to cell cultures in vitro. Previous studies using Ficoll or dextran as crowders demonstrate that the expression of collagen I and fibronectin in WI38 and WS-1 cell lines are significantly enhanced using the MMC technique. However, this technique has not been validated in primary hypertrophic scar-derived human skin fibroblasts (hHSFs). As hypertrophic scarring arises from the excessive deposition of collagen, this protocol aims to construct a collagen-rich in vitro hypertrophic scar model by applying the MMC technique with hHSFs. This optimized MMC model has been shown to possess more similarities with in vivo scar tissue compared to traditional 2-dimensional (2-D) cell culture systems. In addition, it is cost-effective, time-efficient, and ethically desirable compared to animal models. Therefore, the optimized model reported here offers an advanced &#8220;in vivo-like&#8221; model for hypertrophic scar-related studies.

This protocol describes the use of macromolecular crowding to create an in vitro human hypertrophic scar tissue model that resembles in vivo conditions. When cultivated in a crowded macromolecular environment, human skin fibroblasts exhibit phenotypes, biochemistry, physiology, and functional characteristics resembling scar tissue.

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol ...

Using Magnetometry to Monitor Cellular Incorporation and Subsequent Biodegradation of Chemically Synthetized Iron Oxide Nanoparticles

Magnetic nanoparticles, made of iron oxide, present a peculiar interest for a wide range of biomedical applications for which they are often internalized in cells and then left within. One challenge is to assess their fate in the intracellular environment with reliable and precise methodologies. Herein, we introduce the use of the vibrating sample magnetometer (VSM) to precisely quantify the integrity of magnetic nanoparticles within cells by measuring their magnetic moment. Stem cells are first labeled with two types of magnetic nanoparticles; the nanoparticles have the same core produced via a fast and efficient microwave-based nonaqueous sol gel synthesis and differ in their coating: the commonly used citric acid molecule is compared to polyacrylic acid. The formation of 3D cell-spheroids is then achieved via centrifugation and the magnetic moment of these spheroids is measured at different times with the VSM. The obtained moment is a direct fingerprint of the nanoparticles&#8217; integrity, with decreasing values indicative of a nanoparticle degradation. For both nanoparticles, the magnetic moment decreases over culture time revealing their biodegradation. A protective effect of the polyacrylic acid coating is also shown, when compared to citric acid.

Iron oxide nanoparticles are synthesized via a nonaqueous sol gel procedure and coated with anionic short molecules or polymer. The use of magnetometry for monitoring the incorporation and biotransformations of magnetic nanoparticles inside human stem cells is demonstrated using a vibrating sample magnetometer (VSM).

Magnetic nanoparticles, made of iron oxide, present a peculiar interest for a wide range of biomedical applications for which they are often internalized ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...